I have been trying the inverse PCR method to change a single amino acid, so i designed back to back primers, with the forward primer having the mutation. I then did the PCR, phosphorylation, and ligation protocol (overnight). When I transform by electroporation, there usually is no colonies. When there are colonies (about 2), there are no mutations for the most part. And when there is the desired mutation, there are huge insertions (200 bp) that seem to come from no where. I really dont know what to make of my results, and i would very much appreciate the help. Also if anyone has more troubleshooting techniques, I would be glad to hear.

I assume you are using a good long range proofreading type PCR kit. If not, you should. I would not attempt this with mere Taq polymerase. Especially if the plasmid you are mutating is big. Also you don’t want any A overhangs like you can get with Taq- you want nice blunt ends.

Also, how do you get rid of any leftover template plasmid DNA for your PCR? You do not want that in your transformation. Though that does not seem to be your problem, or you would have too many colonies (unmutated). Are you doing a DpnI digest? How much template DNA do you put in your PCRs?

You might want to consider a site-directed mutagenesis kit such as Invitrogen makes. I’ve had good results from them. Their website has a primer design wizard and I use it-rather than trying to design my own. (I do not work for Invitrogen, or any other supplier.)

Your “huge insertions” might be E. coli genomic, which is always a minor contaminant in plasmid preps. Are the insertions right near your desired mutation in the few colonies that have the mutation? If not, I have another idea for you, but it won’t work if the insertion is right near the desired mutation, alas.

Also, do you do a positive control for your electroporation, to make sure that part of your procedure is OK?

Thank you so much for replying. I was working with Staphylococcus epidermidis, so i cant do a Dpn1 digest. Plasmid template was about 200 ng. I think it was the plasmid size that made it difficult. it was about over 12 kbp.

Ah, that is an important piece of information. I’ve no personal experience with transforming S. epi but I understand the species has a strong restriction/modification system (RM) that rejects “invading” DNA with the incorrect or absent modifications. Can I assume you have one of the mutant strains in which this RM has been disabled? When you PCR-amplify plasmids the product has no modifications even if the template molecule had them, so your amplified plasmid is “foreign” to the S. epi. Did you use a positive control, like some unamplified 12kb plasmid, in your electroporations to see if that worked? That is an important test. Size of the plasmid may be a factor, but you have a lot of other hurdles to overcome.

Does your plasmid have shuttle capability, that is, can you grow it in E. coli as well as S. epi? If so, try making your mutant plasmid in E. coli, then after you confirm that the mutation is present (and you didn’t get any unwanted mutations) by sequencing, then put it back into your S. epi.

Or, looking at a restriction map of your plasmid, can you conveniently cut out a small section containing the sequence you want to modify? If so, try cloning it into some small common vector (pUC, Bluescript, etc.) used for E. coli, then do the modification (either by your method or get a site-directed mutagenesis kit) and transform it into E.coli. Confirm the modification in the new plasmid and then cut out the section and clone it back into your 12 kb plasmid. Can you get the mutated piece of DNA from the clones you made that have "huge insertions" and put it into the original plasmid by cloning?

You can also have a block of DNA synthesized with the mutation in place, and clone that into your plasmid. Several companies that synthesize oligos now offer these synthetic double-stranded fragments and they are getting pretty affordable.Hope something works for you!